Multilayered regulation of autophagy by the Atg1 kinase orchestrates spatial and temporal control of autophagosome formation

Summary Autophagy is a conserved intracellular degradation pathway exerting various cytoprotective and homeostatic functions by using de novo double-membrane vesicle (autophagosome) formation to target a wide range of cytoplasmic material for vacuolar/lysosomal degradation. The Atg1 kinase is one of its key regulators, coordinating a complex signaling program to orchestrate autophagosome formation. Combining in vitro reconstitution and cell-based approaches, we demonstrate that Atg1 is activated by lipidated Atg8 (Atg8-PE), stimulating substrate phosphorylation along the growing autophagosomal membrane. Atg1-dependent phosphorylation of Atg13 triggers Atg1 complex dissociation, enabling rapid turnover of Atg1 complex subunits at the pre-autophagosomal structure (PAS). Moreover, Atg1 recruitment by Atg8-PE self-regulates Atg8-PE levels in the growing autophagosomal membrane by phosphorylating and thus inhibiting the Atg8-specific E2 and E3. Our work uncovers the molecular basis for positive and negative feedback imposed by Atg1 and how opposing phosphorylation and dephosphorylation events underlie the spatiotemporal regulation of autophagy.

In brief Schreiber et al. show that upon autophagy induction, lipidated Atg8 activates the Atg1 kinase. This downregulates Atg8 lipidation as Atg1 phosphorylates two of the enzymes involved in Atg8 lipidation, inhibiting their activity. Phosphorylation of Atg13 further dissociates the Atg1 complex, regulating Atg1 complex dynamics at the phagophore assembly site.

INTRODUCTION
Macroautophagy, hereafter referred to as autophagy, is an intracellular degradation pathway that uses de novo double-membrane vesicle (autophagosome) formation to engulf cytoplasmic material. Degradation of the enclosed cellular material by fusion with the vacuole/lysosome allows recycling of cellular building blocks, explaining the function of autophagy in cellular homeostasis, particularly during nutrient starvation. While bulk autophagy mediates the seemingly random uptake of cytoplasmic material, selective autophagy pathways target potentially cytotoxic structures such as damaged organelles, protein aggregates, or invading pathogens in a highly discriminating manner. Deregulation of autophagy is involved in the etiology of diverse human diseases such as cancer, neurodegeneration, and infectious diseases (Dikic and Elazar, 2018). Hence, modulating this cellular self-defense pathway holds promise for treating some of the most prevalent human diseases and for slowing the cellular aging process.
Covalent conjugation of the ubiquitin-like (Ubl) protein Atg8 to phosphatidylethanolamine (PE) is a hallmark of autophagy. Atg8
Here, we recombinantly expressed and purified the core autophagy machinery, and characterized enzymatic activities and protein-protein interactions directly regulated by Atg1 kinase activity. We show that Atg1 is recruited and activated by Atg8-PE. Surprisingly, Atg1 downregulates Atg8 lipidation by inhibiting the Atg8-specific E2 (Atg3) and E3 (Atg12-Atg5-Atg16), generating negative feedback that self-regulates Atg8-PE levels in the growing phagophore. Moreover, Atg1 kinase activity regulates its own assembly state by triggering the disassembly of Atg1based complexes at the PAS. Our studies thus provide mechanistic insights into Atg1 activation, Atg1 complex dynamics, and the spatiotemporal regulation of Atg8 lipidation, highlighting the phagophore as a critical signaling platform.
In vitro kinase assays confirmed that wild-type (WT) Atg1 readily phosphorylates itself (Figures 1C and S1B) and previously reported in vivo substrates, including Atg2, Atg4, Atg6, Atg9, Atg13, Atg29, and Atg31 (Figures 1C, S1B-S1G, and A C B Figure 1. Reconstitution of the core autophagy machinery allows Atg1 substrate identification in vitro (A) Overview of the S. cerevisiae core autophagy machinery and its respective functions. Autophagosome formation is initiated at the PAS, requiring both Atg1 kinase and PP activity. (B) SDS-PAGE analysis of recombinant Atg proteins expressed and purified for this study. Asterisks, co-purifying insect cell proteins. (C) Autoradiograph of in vitro kinase assays depicting Atg1 substrates. ll OPEN ACCESS S1J). Recombinant Atg1 also phosphorylated substrates so far only observed in mammalian cells (Vps34 and Atg16) (Figures 1C, S1F, and S1G) and Atg proteins not previously described as Atg1 targets, including Atg18, Atg38, and several members of the Atg8 lipidation machinery such as Atg3, Atg12, Atg5, and Atg8 itself (Figures 1C, S1F-S1H, S1K, and S1L). In contrast, Atg7, Atg10, Atg14, and Atg17 were not significantly phosphorylated in vitro ( Figures 1C, S1E, S1G, and S1J), suggesting that they are unlikely Atg1 targets.
To further analyze the specificity of our in vitro system, we compared the phosphorylation of a subset of Atg1 targets with two other protein kinases, Hrr25 and Tpk1 (Figures S1I-S1M), as both have previously been shown to phosphorylate Atg proteins (Stephan et al., 2009;Tanaka et al., 2014;Mochida et al., 2014;Pfaffenwimmer et al., 2014). As expected, Tpk1 also phosphorylated Atg13 ( Figure S1J), while neither Tpk1 nor Hrr25 significantly phosphorylated Atg1 substrates involved in Atg8 lipidation (Figures S1K and S1L). These data suggest that our recombinant system provides a versatile and specific tool to rapidly screen for direct Atg1 substrates, in particular for Atg proteins whose phosphorylation is difficult to detect in vivo due to their inherently low expression levels (Ho et al., 2018).
Atg1 directly binds Atg8 via an AIM (Kraft et al., 2012;Nakatogawa et al., 2012b), which is thought to direct Atg1 to growing phagophores (Suzuki et al., 2013). Mass spectrometry analysis of autophosphorylated Atg1 identified three phosphorylation sites N-terminal to this AIM ( Figure 2E; Table S1), and this region was also phosphorylated in vivo (Hu et al., 2019;Lanz et al., 2021). However, mutating these phosphorylation sites did not aggrevate the bulk autophagy defect of an Atg1 AIM mutant (Figure S2F). To understand whether the Atg1 AIM is required for Atg8-PE-dependent stimulation of substrate phosphorylation, we carried out competition assays monitoring Atg1-dependent substrate phosphorylation in the presence of WT and phosphomimicking AIM peptides ( Figure 2E). Interestingly, Atg3 phosphorylation was strongly impaired by either AIM peptide (Figures 2E and 2F), implying that AIM-dependent Atg8-PE binding is required to stimulate Atg1 kinase activity.
To corroborate this finding, we compared Atg8 and Atg8-PEdependent substrate phosphorylation by Atg1 to an Atg1 AIM mutant with the three proceeding phosphorylation sites mutated to alanine (Atg1 DAIM/S>A ). Surprisingly, the Atg1 DAIM/S>A mutant had no defect when analyzing Atg3 and Atg8-PE phosphorylation in the presence of Atg8-PE ( Figures 2G and 2H).
To understand whether Atg1 DAIM/S>A lost its ability to bind Atg8, we studied the phagophore localization of Atg1 DAIM/S>A in atg11D Figure 2. Atg8-PE stimulates Atg1 kinase activity (A) Atg1-mediated phosphorylation of catalytically inactive Atg1 D211A was monitored in the presence of either 4 mM lipidated Atg8 DR117 (Atg8 DR117 -PE) or nonlipidated Atg8 DG116/DR117 (upper panel). Before the addition of Atg1, Atg8 DR117 and Atg8 DG116/DR117 were incubated with Atg7, Atg3, and liposomes (55% PE, 35% phosphatidylcholine [PC], and 10% phosphatidylinositol [PI]) to promote Atg8 DR117 lipidation. Atg1 D211A (2.5 mM) was added, and in vitro kinase assays were started by the addition of WT Atg1 (50 nM). Control reactions (lower panel) did not contain Atg1 D211A reporting only on Atg1 autophosphorylation. Time points were analyzed by SDS-PAGE and autoradiography. (B) Atg1-dependent phosphorylation of Atg8 and Atg3 was monitored in a time-dependent manner in the presence of either Atg8 (Atg8 DR117 ) or Atg8-PE (Atg8 DR117 -PE). Atg8 was lipidated before starting the in vitro kinase assays by addition of pre-phosphorylated Atg1. Control reactions lacked either Atg7 or Atg8. Time points were analyzed by SDS-PAGE and autoradiography. Asterisk, Atg8-Atg3 conjugate. (C) Relative quantification of Atg1-dependent Atg3 and Atg8/Atg8-PE phosphorylation rates (n = 3) in the presence of Atg8 (by omitting either Atg7 or liposomes) or Atg8-PE. Atg3 phosphorylation rates were also quantified in the absence of Atg8. Representative autoradiographs used for quantification are shown in Figure S2A. (D) Relative rate increase of Atg1-dependent substrate phosphorylation in the presence of Atg8-PE. Atg8-PE-dependent phosphorylation rates were compared to control reactions devoid of Atg7 or liposomes. Relative phosphorylation rates for the indicated Atg proteins were determined using Atg8, Atg3, Atg12-Atg5-Atg16, Vps34 Atg14/Atg38 , and Atg4 C147A as substrates. Atg1 was phosphorylated before substrate addition. (E) Schematic representation of S. cerevisiae Atg1 highlighting the Atg8-interacting motif (AIM) (salmon), kinase, and MIT domains (purple and green, respectively). Atg1 autophosphorylation sites and corresponding phosphomimicking mutations are shown in red. Atg1-dependent phosphorylation of Atg3 was assayed in the presence of WT (AIM WT ) or mutated AIM peptides (AIM AA and AIM WT/S>D ). Competition experiments were carried out in the presence of either Atg8 (bottom row) or Atg8-PE (top row). Longer time points were used to monitor Atg3 phosphorylation in the presence of Atg8. (F) Quantification of the relative Atg1-dependent Atg3 phosphorylation rates in arbitrary units (a.u.) in the presence of the indicated AIM peptides and either Atg8 or Atg8-PE. (G) Phosphorylation of Atg3 and Atg8/Atg8-PE was compared between WT Atg1 and Atg1 DAIM/S>A (Atg1 S418A/S421A/S424A/Y429A/V432A ) in the presence of either Atg8 or Atg8-PE. Different time points are shown for Atg8 and Atg8-PE-containing reactions.
To map its location, we expressed and purified different Atg1 DAIM/S>A truncation mutants ( Figure S2J). While the largely disordered central region harbored no additional AIM (Figure S2K), both the N-terminal kinase and the C-terminal microtubule-interacting and transport (MIT) domains weakly bound Atg8. However, as Atg8 binding did not significantly increase upon autophosphorylation (Figures S2K and S2L) and no linear AIM could be identified so far, a more complex mechanism involving both kinase and MIT domains is conceivable.
We conclude, therefore, that autophosphorylation exposes an additional phosphorylation-regulated AIM in Atg1 and that Atg8-PE dependent recruitment of Atg1 upregulates its kinase activity, stimulating both Atg1 autophosphorylation and phagophoreassociated substrate phosphorylation.
To understand whether Atg1 needs to be part of the complex to trigger disassembly, we added sub-stoichiometric amounts of WT Atg1 to a preformed, catalytically inactive Atg1 D211A complex. Strikingly, the addition of WT Atg1 dissociated an otherwise stable complex, demonstrating that phosphorylation in trans is sufficient to trigger complex disassembly ( Figure S3E). Notably, Tpk1 was unable to destabilize inactive Atg1 complexes ( Figures  S3F and S3G), although it phosphorylated Atg13 in vitro (Figure S1J) and in vivo (Stephan et al., 2009).
To test whether Atg1 kinase activity also affects Atg1 complex dynamics in vivo, we carried out fluorescence recovery after photobleaching (FRAP) experiments. Atg13-neonGreen fluorescence rapidly recovered at the PAS in atg11D cells expressing WT Atg1, but not in cells expressing Atg1 D211A ( Figure 3D). Moreover, Atg13 and all other Atg1 complex subunits strongly accumulated at the PAS in starved atg1 D211A cells (Figures 3E and 3F). This accumulation was not caused by a general block in autophagy, as deletion of Atg8 or Atg9 had no effect ( Figure 3E). Notably, Atg13 PAS accumulation was also detected in atg1 D211A cells grown in nutrient-rich medium ( Figure 3E), suggesting that kinase activity may also regulate Atg1 complex dynamics during selective autophagy.
To investigate the underlying mechanism of Atg1 complex disassembly, we studied how the phosphorylation of all three dissociation products affects their subunit-subunit interactions. While phosphorylation of Atg1 and Atg17-Atg29-Atg31 did not (G) Atg13 was phosphorylated using substoichiometric amounts of Atg1. ATP was depleted using apyrase and phosphorylated, and non-phosphorylated Atg13 was immobilized. Autophosphorylated Atg1, Atg1 D211A , or Atg17-Atg29-Atg31 were added, and Atg13 and co-purifying proteins were eluted and analyzed by SDS-PAGE. Asterisk, Atg13 truncation. (H) Serines and threonines mutated to alanine in the Atg13 S/T>A mutant are shown with respect to the N-terminal HORMA domain, Atg17 binding sites, and MIT-interacting motifs (MIMs). In vivo phosphorylation sites are marked by asterisks with putative Atg1-dependent phosphorylation sites underlined. Serines highlighted in black are phosphorylated by recombinant Atg1 in vitro. (I) Atg13 WT or Atg13 S/T>A were mixed with Atg1 and Atg17-Atg29-Atg31 (I) and incubated with either ATP/Mg 2+ or l-PP. Atg13 and Atg13 S/T>A were immobilized and elutions (E) were analyzed by SDS-PAGE. (J) WT and atg11D cells expressing either Atg13 WT or Atg13 S/T>A were nitrogen starved (7 h), and processing of precursor Ape1 (prApe1) to its mature form (mApe1) was monitored by western blotting (WB). (K) Pho8D60 assay measuring bulk autophagy in atg1D strains and yeast expressing either Atg13 WT or Atg13 S/T>A . Cells were either exponentially grown in YPD medium or nitrogen starved for 2, 4, 7, or 18 h. Alkaline phosphatase activity was measured (n = 3) and plotted as relative Pho8D60 activity with standard deviation. (L) The indicated yeast strains were exponentially grown in YPD medium and Cvt pathway-dependent Ape1 processing was monitored by WB. (M) Model summarizing phosphorylation-dependent dissociation of the Atg1 complex.
Atg13 contains a HORMA domain in its N terminus that may act as a phosphorylation sensor (Jao et al., 2013). However, an Atg1 complex lacking the HORMA domain had no obvious disassembly defects ( Figure S3I). Likewise, an Atg1 complex containing an Atg13 truncation mutant lacking both the N-terminal HORMA domain and the C-terminal region (Atg13 DHORMA_DC ) could still dissociate upon Atg1-mediated phosphorylation regardless of whether we mutated the three TORC1 phosphorylation sites in the Atg17-Atg13 interface (Atg13 DHORMA_DC_TOR_S>A ) (Figures 3H and S3J-S3L). Importantly, mass spectrometry analysis of Atg13 detected multiple phosphorylation sites in this central region that are phosphorylated by Atg1 in vitro ( Figure 3H; Table S1), with most of them also phosphorylated in vivo ( Figure 3H) (Fujioka et al., 2014;Hu et al., 2019;Lanz et al., 2021). Mutating the main Atg1-dependent phosphorylation sites outside the Atg1 binding region (Atg13 DHORMA_DC_pS>A ) ( Figure S3J) was not sufficient to stabilize the Atg1 complex ( Figure S3L). As this Atg13 mutant still underwent an electrophoretic mobility shift upon Atg1-mediated phosphorylation ( Figure S3L), we mutated all of the serines and threonines within this region to alanine (Atg13 DHORMA_DC_S/T>A ), stabilizing the resultant complex in the presence of ATP/Mg 2+ ( Figure S3M). Next, we assembled an Atg1 complex with the same set of mutations in full-length Atg13 (Atg13 S/T>A ). While the resulting complex was still able to eject Atg17-Atg29-Atg31, Atg1 remained bound to Atg13 S/T>A even in the presence of ATP/Mg 2+ ( Figures 3I, S4A, and S4B), establishing that phosphorylation of the Atg13 central region by Atg1 specifically regulates Atg1-Atg13 dissociation. Notably, we did not detect significant changes in substrate phosphorylation between Atg1, Atg1-Atg13, and Atg1-Atg13 S/T>A ( Figure S4C), demonstrating that Atg13 S/T>A binding does not change Atg1 kinase activity.
Consistent with its ability to support Atg1 complex assembly ( Figures 3I, S4A, and S4B), Atg13 S/T>A formed a dot-like structure adjacent to the vacuole similar to WT Atg13 ( Figure S4D). Likewise, both Atg13 WT and Atg13 S/T>A accumulated at the PAS in cells expressing catalytically inactive Atg1 D211A (Figure S4D). Atg13 S/T>A , however, did not accumulate at the PAS in WT cells ( Figure S4D), as Atg13 PAS recruitment depends on Atg17 (Suzuki et al., 2007;Cheong et al., 2008) ( Figure S4E) and Atg13 S/T>A dissociation from Atg17 was largely unimpaired ( Figures 3I and S4A). As a consequence, neither Atg17 nor Atg1 accumulated at the PAS upon starvation in cells expressing Atg13 S/T>A ( Figure S4F). These data suggest that Atg13 S/T>A promotes assembly of an active Atg1 complex at the PAS, which is specifically impaired in the phosphorylation-mediated dissociation of the Atg1-Atg13 S/T>A subcomplex.
To examine the physiological importance of Atg1-Atg13 complex dissociation, we studied bulk and selective autophagy in Atg13 S/T>A -expressing cells. Strikingly, bulk autophagy was strongly impaired in atg13 S/T>A cells, while the selective cytoplasm-to-vacuole targeting (Cvt) pathway was largely unaltered ( Figures 3J-3L and S4G).
These results demonstrate that the Atg1 complex is a highly dynamic entity that needs to cycle between an assembled and a disassembled state, driven at least in part by Atg13 phosphorylation and counteracting dephosphorylation ( Figure 3M).

Atg1-dependent phosphorylation of Atg12 inhibits E3 activity
To dissect the mechanism of this inhibition, we individually phosphorylated the different Atg1 targets. We purified Atg1 phosphorylated Atg12-Atg5-Atg16, allowing us to carry out Atg8 lipidation assays in the absence of Atg1. Atg8 lipidation was delayed when phosphorylated but not dephosphorylated E3 was added (Figures 4C and S5G), suggesting that Atg1 directly inhibits the E3. The Atg5-Atg16 complex is not active as E3 unless conjugated to Atg12 (Hanada et al., 2007;Metlagel et al., 2013). However, neither E3 formation nor stability were affected by Atg1 kinase activity ( Figures S5A and S5B). As Atg12 and Atg12-Atg5 are both Atg1 targets ( Figure 1C), we investigated the role of Atg1-dependent Atg12 phosphorylation. We purified endogenous Atg12 from nitrogen-starved yeast expressing either WT Atg1 or catalytically inactive Atg1 D211A and compared the electrophoretic mobility of the Atg12-Atg5 conjugate by Phos-tag SDS-PAGE. Only the Atg12-Atg5 conjugate purified from WT cells migrated as two distinct bands, with a unique slower migrating band consistent with Atg1-dependent Atg12 phosphorylation in vivo ( Figure 4D). Mass spectrometry analysis of in vitro phosphorylated Atg12 identified 12 Atg1 phosphorylation sites in the N-terminal extension and 4 in the Ubl domain ( Figure 4E; Table S1). Consistently, mutation of these phosphorylation sites strongly reduced the (B) Recombinant Atg3, Atg7, and Atg12-Atg5-Atg16 were incubated with WT Atg1, Atg1 D211A , or l-PP in the presence of ATP/Mg 2+ and PP inhibitors. Liposomes were added and lipidation reactions were started by the addition of Atg8 (Atg8 DR117 ) or Atg8 N (Atg8 S3A/T4A/S7A/DR117 ). Samples were taken at indicated time points and analyzed by urea-SDS-PAGE and Sypro Ruby staining. (C) Atg8 lipidation was monitored in the presence or absence of E3 and in the presence of an E3 pre-phosphorylated by Atg1 (pE3). Lipidation reactions were set up as illustrated in Figure S5G. Samples were taken and analyzed as in (B). The average lipidation ± standard deviation is plotted for each time point (n = 3; bottom panel). (D) Endogenous SF-tagged Atg12 was purified from nitrogen-starved yeast expressing WT Atg1 or Atg1 D211A . The electrophoretic mobility of the Atg12-Atg5 conjugate was compared by Phos-tag SDS-PAGE and WB. (E) Schematic overview of S. cerevisiae Atg12 highlighting the N-terminal extension, Ubl domain, and Atg1-dependent in vitro phosphorylation sites. Asterisks indicate phosphorylation sites also identified in vivo (Hu et al., 2019;Lanz et al., 2021). Underlined sites were previously reported to be phosphorylated by Atg1 in vitro (Hu et al., 2019). (F) Bulk autophagy was quantified in Atg12-, Atg12 S>A_ALL -, Atg12 S>D_ALL -, and Atg12 S>D_Ubl -expressing or atg12D cells using the Pho8D60 assay. Cells were exponentially grown in YPD medium or nitrogen starved for 2.5, 5, or 7.5 h. (G) Bulk autophagy was monitored in nitrogen-starved atg11D cells expressing WT Atg12, Atg12 S>A_ALL , Atg12 S>D_ALL , or Atg12 S>D_Ubl . Ape1 processing was monitored by WB. (B) Atg3, Atg7, and Atg8 were individually incubated with WT Atg1 or Atg1 D211A and ATP/Mg 2+ before combining to promote Atg3 charging. Liposomes were added to monitor Atg8 discharge in a time-dependent manner. Samples were analyzed by SDS-PAGE under reducing or non-reducing conditions. (C) Atg8 lipidation assays were carried out using WT Atg3, Atg1-phosphorylated Atg3 (pAtg3), or l-PP or PP2A Rts1 -treated pAtg3. Samples were taken at the indicated time points, and Atg8 lipidation was analyzed by urea-SDS-PAGE and Sypro Ruby staining.
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Atg1-mediated phosphorylation of Atg12 ( Figure S5H), confirming that we identified the majority of the Atg1-dependent phosphorylation sites.
To test their functional significance, we compared bulk and selective autophagy in cells expressing phosphorylation-deficient and phosphomimicking mutants, Atg12 S>A_ALL and Atg12 S>D_ALL , respectively. While Atg12 S>A_ALL -expressing cells showed only a mild bulk autophagy defect (Figures 4F and 4G) and a fully functional Cvt pathway ( Figure  To corroborate these findings, we assembled an E3 complex containing the Atg12 S>D_Ubl mutant in vitro. Although Atg12 S>D_Ubl was efficiently conjugated to Atg5, allowing the formation of a phosphomimicking E3 ( Figure S5M), the resultant E3 failed to stimulate Atg8 lipidation ( Figure 4I). We therefore conclude that the phosphorylation of the Ubl domain of Atg12 by Atg1 efficiently blocks Atg8 lipidation, explaining at least in part the strong autophagy and Atg8 lipidation defect observed in Atg12 S>D_Ubl -expressing cells.

Atg1-mediated phosphorylation of Atg3 inhibits Atg8 lipidation
To examine whether E3 phosphorylation is the sole reason for the observed inhibition of Atg8 lipidation, we took advantage of the fact that Atg8 lipidation does not require E3 activity in vitro. Strikingly, Atg1-mediated phosphorylation efficiently blocked Atg8 lipidation even in the absence of the E3 ( Figure 5A). This inhibition was specific to Atg1, as addition of other kinases (PKA, Tpk1, Hrr25, and Plk1) did not alter Atg8 lipidation ( Figure S6A). Atg1 kinase activity did not affect the formation of the thioesterlinked Atg3-Atg8 intermediate ( Figure S6B), demonstrating that Atg1 neither regulates E1 activity nor transthiolation nor Atg3 charging. We therefore tested Atg8 discharge to PE-containing liposomes and observed a striking discharge defect in the presence of Atg1 kinase activity ( Figure 5B).
To confirm that Atg3 phosphorylation specifically inhibits Atg8 lipidation, we pre-phosphorylated Atg3 using Atg1 (pAtg3) and further purified it to avoid simultaneous Atg8 and Atg7 phosphorylation in Atg8 lipidation assays. Strikingly, pAtg3 was not able to lipidate Atg8, and this effect could be reversed by incubating pAtg3 with either l-PP or PP2A Rts1 ( Figure 5C). To further investigate the molecular mechanism of Atg3 inhibition, we mapped the Atg1-dependent phosphorylation sites in vitro and in vivo.
The N terminus of Atg3 comprises an amphipathic alpha helix ( Figures 5D and 5E), which mediates membrane binding (Hanada et al., 2009;Hervá s et al., 2017;Nath et al., 2014). The N-terminal Atg1 phosphorylation sites map largely to the hydrophilic face of the amphipathic helix ( Figure 5E), making them unlikely to directly interfere with membrane binding. Atg1dependent phosphorylation did not significantly change the binding of Atg3 to Atg8-PE-containing liposomes ( Figure S6M). Notably, the N terminus of Atg3 is positioned in close proximity to the active site ( Figures 5D and S6N). Adjacent to the active site is a highly conserved, positively charged canyon that coordinates two sulfate ions in the Atg3 crystal structure (Yamada et al., 2007) (Figures 5D and S6N-S6Q), which can suggest the binding of phosphoryl groups. A phosphorylated N terminus could provide a high density of negatively charged phosphoryl groups to engage with the positively charged canyon, thereby blocking the active site. Consistent with such a model, the (D) Schematic domain overview and surface representation of S. cerevisiae Atg3 (PDB: 2DYT) highlighting phosphorylation sites and unique functional elements, including the N-terminal amphipathic helix (green), the E1, E2, and E3 interacting region (E123IR), the active site cysteine, and the AIM. Sites phosphorylated in vivo and by Atg1 in vitro are highlighted in bold black. In vivo phosphorylation sites regulated by Atg1 are underlined. Phosphorylation sites detected either in vitro or in vivo are shown in gray and black, respectively. (E) Sequence alignment of the Atg3 N terminus with Atg1-dependent in vitro and in vivo phosphorylation sites colored in red and orange. Residues in yellow indicate potential phosphorylation sites in other organisms. The helical wheel representation of the N-terminal amphipathic helix is shown below. (F) Atg8 N lipidation was compared in the presence or absence of Atg1 using either WT Atg3, Atg3 S4A/S7A/S8A , or Atg3 S4A/T5A/S7A/S8A . Atg7, Atg3, and Atg8 N were separately incubated with or without Atg1 before starting the lipidation reaction. Samples were analyzed by urea-SDS-PAGE and Sypro Ruby staining. (G) Atg8 lipidation was studied in the presence of Atg3 or Atg3 S4D/T5E/S7D/S8D . In a separate reaction, the E3 was added to Atg3 S4D/T5E/S7D/S8D -containing reactions. Atg8 lipidation was analyzed as in (F). (H) Bulk autophagy was quantified in nitrogen-starved (4 h) WT Atg3, atg3D, and Atg3 S4D/T5E/S7D/S8D -expressing cells using the Pho8D60 assay (n = 3). (I) The Pho8D60 assay was used to quantify bulk autophagy in yeast co-expressing Atg3 S4A/T5A/S7A/S8A with either Atg12 S>A_ALL or Atg12 S>A_Ubl (n = 3). presence of negatively charged peptides is sufficient to inhibit Atg8 lipidation in vitro ( Figure S6R). Glycine 26, which is invariant among Atg3 orthologs ( Figures 5E and S6N), could act as a hinge to provide the N terminus with enough flexibility to mediate binding to the basic canyon. Rigidifying this pivot by replacing the highly conserved glycine with a proline completely abolished Atg3 activity ( Figure S6H). These data suggest that Atg1-mediated phosphorylation of the Atg3 N terminus induces a conformational change that blocks the E2 active site, and as a result, inhibits Atg8 discharge.

DISCUSSION
To better understand the complex role of Atg1 in orchestrating autophagy, we combined in vitro reconstitution with cell-based assays to systematically study the functional consequences of Atg1-mediated phosphorylation. We find that Atg1 autophosphorylation and substrate phosphorylation are both highly upregulated by Atg8-PE. Atg1 autophosphorylation stimulates membrane recruitment of Atg1 and promotes the phosphorylation of phagophore-associated Atg1 substrates. Surprisingly, Atg1 inhibits Atg8 lipidation by phosphorylating the Atg8-specific E2 and E3, thereby limiting the Atg8-PE density in the growing phagophore. Atg1 recruitment by Atg8-PE therefore establishes both positive and negative feedback. Finally, Atg1-mediated phosphorylation of Atg13 triggers rapid disassembly of Atg1-based complexes at the PAS, resulting in the dissociation of Atg17-Atg29-Atg31 and Atg1 from Atg13, with the latter playing an important role during bulk autophagy. Since Atg1 complex disassembly and inhibition of Atg8 lipidation can be reversed by PP activity, we conclude that autophagy is driven by the spatiotemporal regulation of phosphorylation and counteracting dephosphorylation events.
Atg1 complex formation appears to be spatiotemporally regulated as Atg13 and Atg17 only localize to the PAS and are absent from growing autophagosomal membranes, despite Atg8-PEdependent recruitment of Atg1 (Suzuki et al., 2013). We show that Atg1 complexes at the PAS are intrinsically unstable due to Atg1-mediated phosphorylation of Atg13, which triggers its dissociation from Atg1 and Atg17-Atg29-Atg31. Despite the overall desphosphorylation of Atg13 in response to nutrient starvation (Kamada et al., 2000), our data suggest that Atg1 continuously phosphorylates Atg13 and thus destabilizes the Atg1 complex even in the absence of TORC1-mediated phosphorylation. PAS localized PPs dephosphorylating Atg13 (Memisoglu et al., 2019) are thus likely to drive the rapid reassembly of Atg1-based complexes. It is tempting to speculate that the resultant Atg1 complex dynamics not only provide the molecular basis for the liquid-like properties of the PAS (Fujioka et al., 2020) but also drive autophagosomal membrane formation. While Atg1 complex assembly is thought to tether Atg9-containing vesicles (Rao et al., 2016), dissociation of the Atg1 complex may promote the release of membrane fusion intermediates, freeing up Atg9 binding sites to drive rapid phagophore expansion aided by Atg2-dependent lipid transfer (Matoba et al., 2020;Osawa et al., 2019;Valverde et al., 2019;Maeda et al., 2019). The ejection of Atg17-Atg29-Atg31 may also shape autophagosomal membranes (Bahrami et al., 2017) or help terminate autophagy by continuously exposing the TORC1 phosphoryla-tion sites in Atg13 (Fujioka et al., 2014;Yamamoto et al., 2016), thereby allowing rapid sensing of TORC1 reactivation upon nutrient replenishment.
Mutational analysis revealed the functional significance of Atg1 complex dissociation, as cells expressing an Atg13 mutant unable to dissociate from Atg1 are largely defective for bulk autophagy. As these cells are proficient in forming smaller Cvt vesicles under nutrient-rich conditions, our work highlights further mechanistic differences between bulk and selective autophagy. Given the strong conservation of Atg1 and Atg13, the described oscillatory behavior of human Atg13 during mitophagy (Dalle Pezze et al., 2021), and the accumulation of Ulk1 in the presence of Ulk1 inhibitors (Petherick et al., 2015), it is likely that similar mechanisms also regulate Atg1 complex dynamics in higher eukaryotes.
In addition to Atg1 complex dynamics and autoactivation (Kijanska et al., 2010;Yeh et al., 2010) Atg1 phosphorylation also stimulates its own recruitment to growing autophagosomal membranes by exposing at least one additional phosphorylation-regulated AIM. Atg8-PE binding further upregulates Atg1 autophosphorylation and substrate phosphorylation. The resultant high-affinity/high-activity state leads to the phosphorylation, recruitment, and activation of other Atg1 molecules, creating a zone of high Atg1 kinase activity along the growing phagophore. A concomitant decrease in PP activity away from the PAS, as judged by the inability of phagophore-bound Atg1 to (re)associate with Atg13 and Atg17 (Suzuki et al., 2013), may further enhance this positive feedback. While Atg1 activation at the PAS requires Atg13-and Atg17-dependent clustering of Atg1 complexes (Yamamoto et al., 2016), Atg8-PE-induced activation of Atg1 provides a means to sustain high Atg1 kinase activity along growing phagophores in the absence of Atg13 and Atg17. Most Atg1 substrates are membrane associated, containing either AIMs or other membrane-targeting regions (Baskaran et al., 2012;Birgisdottir et al., 2019;Bozic et al., 2019;Herman et al., 1991;Kaufmann et al., 2014;Ngu et al., 2015;Romanov et al., 2012;Rostislavleva et al., 2015;Suzuki et al., 2013). Hence, Atg8-PE-containing membranes act as scaffolds to recruit both Atg1 and its substrates. This increases their local concentrations and restrains diffusion to a two-dimensional surface, thereby increasing substrate phosphorylation along the growing autophagosomal membrane. This mechanism is distinct from the initial stages of Atg1 activation during selective autophagy, when the cargo itself clusters and activates Atg1 with the help of cargo adaptors and the selectivity factor Atg11 (Kamber et al., 2015). Nevertheless, it is conceivable that Atg8-PE-dependent Atg1 activation also contributes to Atg1 activation during selective autophagy, but akin to bulk autophagy, this activation mechanism would act downstream of Atg8 lipidation.
Atg8-PE not only recruits Atg1 but also the Atg8-specific E2 and E3 to growing phagophores (Ngu et al., 2015;Suzuki et al., 2013). Surprisingly, we find that Atg1-mediated phosphorylation downregulates their enzymatic activities. This Atg1-mediated negative feedback is functionally important in vivo, since bulk autophagy is strongly impaired in cells co-expressing phosphorylation-deficient Atg3 and Atg12 mutants. High levels of Atg8-PE have been shown to affect membrane morphology, resulting in local membrane deformations (Knorr et al., 2014). Hence, self-limiting Atg8-PE levels in the forming autophagosomal ll OPEN ACCESS Article membrane by phosphorylating two of the key enzymes may provide a means to control phagophore expansion, preventing membrane deformations. Eukaryotic cells also need to be resourceful with their available Atg8 pool, especially when facing long periods of starvation. Autophagosome formation severely reduces the pool of free Atg8 since a substantial portion of Atg8-PE is degraded in the vacuole together with the inner autophagosomal membrane. Self-limiting Atg8 lipidation may thus reflect a need for keeping a sufficiently large Atg8 pool. Consistently, Atg8 protein levels are upregulated upon bulk autophagy induction (Xie et al., 2008), and mislocalized Atg8-PE is continuously retrieved from non-autophagosomal membranes by an Atg4-dependent proofreading mechanism replenishing soluble Atg8 (Nakatogawa et al., 2012a).
While E3 inhibition is at least in part mediated by Atg1-mediated phosphorylation of the Atg12 Ubl domain, Atg3 inhibition is mediated by phosphorylation of its N-terminal extension, preventing Atg8 discharge to PE. Our data suggest a model in which phosphorylation of the flexible N terminus blocks the Atg3 active site by promoting an electrostatic interaction with the adjacent highly conserved basic canyon. Atg1-mediated inhibition of Atg3 is distinct from the recently described Atg3 auto-inhibition, which uses the E1, E2, and E3 interacting region to restrain the conformation of the catalytic loop in Atg3 in a phosphorylationindependent manner (Zheng et al., 2019). This E1, E2, and E3 interacting region-dependent auto-inhibition protects from spurious discharge to non-autophagosomal membranes since it couples Atg8 lipidation to E3 binding. In contrast, Atg1-mediated inhibition of Atg3 cannot be reversed by the E3, requiring instead PP activity. Atg1 can thus inhibit Atg8 lipidation, even in the presence of the E3, and therefore regulate Atg8 lipidation along the growing phagophore where Atg3 and the E3 colocalize (Ngu et al., 2015;Suzuki et al., 2013).
Our work emphasizes how in vitro reconstitution approaches can help us understand complex intracellular signaling networks at the molecular level. It paves the way for more detailed mechanistic studies investigating the spatiotemporal regulation of bulk and selective autophagy by both protein kinase and phosphatase signaling.
Limitations of the study While our results clearly show that the Atg1 complex is a highly dynamic entity, mediated at least in part by Atg1-dependent Atg13 phosphorylation and counteracting dephosphorylation, further work is required to understand why its continuous disso-ciation and reassembly is required for bulk autophagy but dispensible for selective autophagy.
Moreover, although we are starting to understand the physiological importance of self-regulating Atg8-PE levels, more advanced microscopy experiments are needed to visualize the morphological consequences of unbalanced Atg8 lipidation in cells.
Finally, although co-expressing the phosphorylation-deficient Atg3 S4A/T5A/S7A/S8A and Atg12 S>A_ALL mutants results in a strong bulk autophagy reduction, it is likely that fully disabling Atg1mediated E2 and E3 inhibition will exhibit an even more pronounced autophagy defect, aiding future studies. Identifying an Atg1 mutant that can no longer interact with Atg8-PE regardless of its phosphorylation status may be an alternative means to study autophagy in cells unable to self-limit Atg8 lipidation and to trigger positive feedback. However, we still lack a catalytically active Atg1 mutant that has fully lost its ability to bind Atg8-PE, regardless of its phosphorylation status. The previously characterized Atg1 AIM mutant is likely to underestimate the physiological relevance of recruiting and activating Atg1 along the growing phagophore, restricting its use.

STAR+METHODS
Detailed methods are provided in the online version of this paper and include the following:

RESOURCE AVAILABILITY
Lead contact Further information and requests for reagents should be directed to and will be fulfilled by the Lead Contact Anne Schreiber (anne. schreiber@crick.ac.uk).

Materials availability
Plasmids generated in this study are available from the Lead Contact without restriction or require a completed Materials Transfer Agreement if there is potential for commercial application.
Data and code availability d The mass spectrometry data have been deposited to the ProteomeXchange Consortium via the PRIDE (Perez-Riverol et al., 2019) partner repository and are publicly available. The accession number is listed in the key resources table. All other data are provided in the manuscript and supplemental information. d This paper does not report original code. d Any additional information required to reanalyze the data reported in this paper is available from the lead contact upon request.

S. cerevisiae strains and media
All yeast strains used in this study are derived from Saccharomyces cerevisiae BY4741 (MATa; his3D1; leu2D0; met15D0; ura3D0) and are summarized in Table S2. Strains were created by transforming the linearized plasmids listed in Table S3. Yeast were grown in YPD (1% yeast extract, 2% peptone, and 2% glucose) or synthetic defined (SD) medium (0.17% yeast nitrogen base, 0.5% ammonium sulfate, 2% glucose and amino acids as required). Starvation experiments were carried out by growing yeast in nitrogen starvation (SD-N) medium (0.17% yeast nitrogen base without amino acids and ammonium sulfate and 2% glucose).

Insect cells and media
Insect cells (Sf9 and High Five cells; Invitrogen) were grown in Sf900 II medium (GIBCO) supplemented with 0.1X Penicillin-Streptomycin-Glutamine (GIBCO).

METHOD DETAILS
Cloning and plasmids All S. cerevisiae genes were PCR amplified from genomic DNA. Restriction sites and tags were introduced by PCR. Mutations and gene fusions were generated by splicing by overlap extension (Heckman and Pease, 2007)

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Plasmids used for yeast strain construction Plasmids used for yeast strain construction are listed in Table S3. All plasmids are pCR-Blunt II or IV TOPO (Invitrogen) derived. Constructs to generate point mutations or gene fusions contained the gene specific promoter ($500 bp upstream of the gene specific start codon) as one region of homology, the mutated open reading frame or gene fusion, the terminator sequence ($150-300 bp downstream of the gene specific stop codon), the selection cassette and a second region of homology downstream of the terminator sequence (300-500 bp). Gene deletions were generated by fully replacing the target gene with the indicated selection cassette. The resultant yeast strains were verified by PCR or sequencing.

Plasmids used for baculovirus generation and insect cell expression
All genes were cloned into the pFBDM transfer plasmid and the resultant plasmids are listed in Table S3. Plasmids were transformed into DH10Multibac cells and bacmids were isolated using isopropanol precipitation (Schreiber et al., 2011).

Protein expression in bacteria
For bacterial protein expression, plasmids listed in Table S3 were transformed into BL21-CodonPlus (DE3)-RIL cells (Agilent) unless stated otherwise. Cells were grown at 37 C in TB medium supplemented with ampicillin (100 mg/ml) and chloramphenicol (25 mg/ml). Cells were grown shaking at 220 rpm until they reached an OD 600 of 0.8. Cells were moved on ice and protein expression was induced with 0.5 mM isopropyl-b-D-1-thiogalactopyranoside (IPTG). Protein expression was carried out overnight at 18 C. Cells were harvested at 4000 rpm for 10 minutes.

Protein expression in insect cells
Bacmids were prepared by isopropanol precipitation. GeneJuice was used to transfect Sf9 cells with bacmids. Viruses were further amplified using standard procedures. All proteins were expressed in High Five insect cells using Sf-900 II SFM medium. Cells were infected with a multiplicity of infection (MOI) greater than 2. Protein expression was carried out at 27 C with cells shaking at 140 rpm. Cells were harvested after three days.

Purification of Atg proteins and protein complexes
If not stated otherwise S. cerevisiae Atg proteins were purified at 4 C. Pre-cooled lysis buffer containing 50 mM Tris HCl pH 8.3, 300 mM NaCl (180 mM NaCl for protein complexes), 5% glycerol, 2 mM DTT, EDTA free protease inhibitor tablets (Roche), 2 mM EDTA, 0.2 mM PMSF, 1 mM benzamidine and Pierce universal nuclease was added to bacterial or insect cell pellets. The lysis buffer used for bacterial protein purifications was supplemented with lysozyme (100 mg/ml). Protease inhibitor tablets and irreversible protease inhibitors were omitted for the purification of enzymes with an active site cysteine. Instead PMSF (0.2 mM) leupeptin (10 mM), pepstatin A (10 mM) and EDTA (4 mM) were used. Cells were lysed by sonication and spun at 20,000 rpm for one hour using a JA-20 rotor.

Affinity purification
Supernatants were loaded onto a StrepTactin column (QIAGEN) or GST trap (GE Healthcare) pre-equilibrated with wash buffer composed of 50 mM Tris HCl pH 8.0, 300 mM NaCl (180 mM NaCl for protein complexes), 5% glycerol and 2 mM DTT. The column was washed with 10 column volumes (CV) wash buffer before proteins were eluted with 5 CV wash buffer containing either 2.5 mM desthiobiotin (StrepTactin) or 10 mM reduced glutathione (GST). Depending on the experiment, tags were cleaved overnight at 4 C using PreScission (3C) protease and a protease to protein molar ratio of 1:50.

Ion exchange chromatography
Protein containing afffinty purification fractions were pooled and diluted to a final salt concentration of 100 mM NaCl and subjected to ion exchange chromatography. Apart from Atg8 all proteins and protein complexes were purified by anion exchange chromatography using a ResQ column (GE Healthcare) applying a salt gradient from 50 to 700 mM NaCl (ResQ buffer base: 20 mM HEPES-NaOH pH 8.0, 5% glycerol and 2 mM DTT). Protein containing fractions were pooled, concentrated and either snap-frozen or further purified/analyzed by size exclusion chromatography. Proteins which were cleaved by PreScission protease were passed back over the initial affinity matrix in order to remove the tag or uncleaved protein.

Size exclusion chromatography
Samples were loaded on a size exclusion chromatography (SEC) column (Superose 6, Superdex 200 or Superdex 75 depending on the size of the protein/protein complex) pre-equilibrated in SEC buffer (20 mM HEPES NaOH pH 7.4, 180 mM NaCl, 5% glycerol and 2 mM DTT). Samples were concentrated using Amicon Ultra concentrators.

Atg8 purification
The affinity purification step was carried out as stated above. Tags were cleaved overnight using PreScission protease. StrepTactin elutions were diluted to a salt concentration of 30 mM NaCl using ResS Buffer A (20 mM MES pH 6.2, 5% glycerol and 2 mM DTT). 500 mM NaCl (ResS buffer base: 20 mM MES pH 6.2, 5% glycerol and 2 mM DTT). The pH of the pooled peak fractions was adjusted to 8.0 and the sample was passed back over a 5 mL StrepTactin column. The StrepTactin column was washed with 1 CV of wash buffer and the flow-through and wash fractions were concentrated using Amicon concentrators (3 kDa cut off). The sample was run on a HiLoad 16/600 Superdex 75 column pre-equilibrated in SEC buffer (20 mM HEPES NaOH pH 7.4, 180 mM NaCl, 5% glycerol, 2 mM DTT). Peak fractions were concentrated and snap frozen.
Atg12-Atg5 was immobilized on a StrepTactin column prior to addition of sonicated bacterial cell lysates containing either Atg16 or Atg16-FLAG. The column was washed with 10 CV of wash buffer (50 mM Tris-HCl pH 8.0, 200 mM NaCl, 5% glycerol and 2 mM DTT) and the resultant E3 complex was eluted using 5 CV of elution buffer (wash buffer supplemented with 2.5 mM desthiobiotin). The E3 was further purified by anion exchange and size exclusion chromatography using a ResQ and Superose 6 column respectively (buffer composition as stated in the general protein purification protocol above). Note: The E3 was loaded on the ResQ column in elution buffer to avoid precipitation. The SEC run was carried out in SEC buffer containing 20 mM HEPES NaOH pH 7.4, 200 mM NaCl, 5% glycerol and 2 mM DTT.

Purification of full-length Atg9
Insect cells expressing full length Atg9 were lysed by passing the cell suspension through an EmulsiFlex. The lysate was cleared using a low speed spin (9000 g for 30 minutes), followed by centrifugation of the resulting supernatant at 40,000 rpm for 1 hour (Ti-45 rotor; Beckman). Pelleted membranes were resuspended in solubilization buffer (50 mM Tris HCl pH 8.0, 300 mM NaCl, 5% glycerol, 2 mM DTT) containing 1% n-Dodecyl-b-D-Maltopyranoside (DDM). The sample was incubated for 4 hours at 4 C before centrifuging at 40,000 rpm for 1 hour (Ti-45 rotor; Beckman). The supernatant was loaded onto a StrepTactin column pre-equilibrated in wash buffer (50 mM Tris HCl pH 8.0, 300 mM NaCl, 2 mM DTT containing either 0.6 mM DDM or 0.6 mM Lauryldimethylamine-N-Oxide (LDAO)).
The column was washed with 10 CV of wash buffer and the protein was eluted using wash buffer containing 2.5 mM desthiobiotin. StrepTactin elutions were concentrated and used directly.

Purification of pre-phosphorylated Atg3
Atg3 was immobilized on a StrepTactin column (QIAGEN) and phosphorylated using recombinant Atg1 (100 nM). Phosphorylation reactions were carried out in 20 mM HEPES-NaOH pH 7.4, 180 mM NaCl, 2 mM ATP, 10 mM MgCl 2 for 4 hours at 30 C. The phosphorylated E2 was eluted in StrepTactin elution buffer (50 mM Tris pH 8.0, 200 mM NaCl, 5% glycerol, 2 mM DTT, 2.5 mM desthiobiotin) and further purified using an anion exchange and size exclusion chromatography step (using a Resource Q and Superdex S75 column respectively). Fractions containing the phosphorylated E2 were pooled and concentrated. For direct control experiments the ''non-phosphorylated'' E2 was purified identically as the pre-phosphorylated E2, however, catalytically inactive Atg1 (Atg1 D211A ) was used instead of wild-type Atg1.
Purification of Tpk1 and Hrr25 kinases SH-SUMO*-Tpk1 was expressed in Rosetta 2(DE3) cells (Novagen) and SH-SUMO*-Hrr25 was expressed in High Five insect cells. Both proteins were purified using a StrepTactin affinity purification step (see ''Purification of Atg Proteins and Protein Complexes''). The SH-SUMO* tag was cleaved off overnight using GST-tagged PreScission protease. Kinases were further purified by SEC using a Superdex 200 column and kinase containing fractions were passed back over a StrepTactin and GST column before concentrating the proteins.

Purification of Sic1
SH-SUMO*-Sic1 was expressed in Rosetta 2(DE3) cells and purified using a StrepTactin affinity purification step (see ''Purification of Atg Proteins and Protein Complexes''). The SH-SUMO* tag was cleaved off overnight using PreScission protease. Sic1 was further purified by ion exchange chromatography using a Resource S column. Protein containing fractions were passed back over a Strep-Tactin column before being concentrated.

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Purification of endogenous S. cerevisiae Atg3 and Atg12 Yeast strains expressing either wild-type or catalytically inactive Atg1 (Atg1 D211A ) and either endogenously SF-tagged Atg3 or Atg12 (yAS_476/yAS_538 and yAS_233/yASC_841 respectively) were grown in YPD medium in a fermenter. Cells were harvested at an OD 600 of $1.0 and washed twice in nitrogen starvation (SD-N) medium. Cells were grown in SD-N medium for another 4 hours before harvesting. Pellets were resuspended in a small volume of resuspension buffer containing 50 mM Tris HCl pH 9.2, 170 mM NaCl, 5% glycerol, 2 mM DTT, 4 mM EDTA and protease and protein phosphatase inhibitors. Cell suspension was frozen in liquid nitrogen before freezer milling. Resuspension buffer supplemented with Pierce Universal Nuclease was added and the resultant lysate was spun at 20,000 rpm for one hour using a JA-20 rotor. Supernatants were loaded onto a pre-equilibrated 1 mL StrepTactin column (QIAGEN). The column was washed with 20 CV of wash buffer (50 mM Tris HCl pH 8.0, 180 mM NaCl, 5% glycerol, 2 mM DTT and protein phosphatase inhibitors). Samples were eluted with BXT buffer (IBA) containing 2 mM DTT (and 7 M UREA when preparing samples for mass spectrometry). Samples were either analyzed by Phos-Tag SDS-PAGE (50 mM Phos-tag acrylamide; 6% polyacrylamide gel; Alpha Laboratories) and Western blot analysis (Atg12) or further processed for phospho-enrichment and subsequent mass spectrometry analysis (Atg3). For the latter biological duplicates were prepared. The SF-tag in Atg3 was inserted between amino acids 266 and 268 as both N-and C-terminal tags impact autophagy (Ngu et al., 2015).
In vitro kinase assays Atg1 was pre-phosphorylated in the presence of 0.5 mM ATP, 2.5 mM MgCl 2 , 1 mg/ml BSA and 0.5X PhosSTOP protein phosphatase inhibitors (Roche). The final Atg1 concentration in the pre-phosphorylation reaction was 1 mM. Pre-phosphorylated Atg1 was diluted 20-fold resulting in a final assay concentration of 50 nM. Substrate phosphorylation was carried out in 20 mM HEPES pH 7.4, 150 mM NaCl, 425 mM ATP, 2.125 mM MgCl 2 , 7.5 mCi [g-32 P]-ATP (3000 Ci/mmol), 0.5 mg/ml BSA and 0.5X PhosSTOP protein phosphatase inhibitors (Roche). Substrates were used at a final assay concentration of 5 mM unless otherwise stated (e.g., the Vps34 Atg14 and Vps34 Atg14/Atg38 complex were used at 2.5 mM). Reactions were started by addition of pre-phosphorylated Atg1. Samples were taken at the indicated time points or after 10 minutes if no time point is specified. Phosphorylation reactions were stopped by addition of 4X SDS sample buffer containing 6 M UREA and 100 mM DTT. Kinase assays containing catalytically inactive Atg1 (Atg1 D211A ), the Vps34 Atg14/Atg38 complex, Hrr25 or Tpk1 were carried out as detailed for wild-type Atg1. Peptides for AIM competition assays were purchased from GenScript, dissolved in DMSO and used at a final concentration of 300 mM. Myelin basic protein (MBP) was purchased from Lucerna-Chem.

Phos-Tag SDS-PAGE
Endogenous SF-tagged Atg12 purified from cells expressing either wild-type Atg1 or catalytically inactive Atg1 D211A was run on a 6% Phos-Tag SDS-PAGE gel containing 50 mM Phos-tag acrylamide (Alpha Laboratories) in 1x Tris glycine running buffer. The gel was washed twice in Protein Transfer Buffer containing 10 mM EDTA and once in EDTA free Transfer Buffer before western blotting using a PVDF membrane. SF-Atg12 was detected using a mouse anti-FLAG M2 antibody (Sigma) and goat anti-mouse IgG (H+L) HRP conjugate (Bio-Rad).
To test the effect of Atg1 mediated phosphorylation on E3 ligase activity, the pre-phosphorylated E3 (pE3) was dephosphorylated using lambda protein phosphatase and PP2A Rts1 . Dephosphorylation was carried out for 2 hours at 30 C. Reactions were stopped by addition of phosphatase inhibitors. Samples were moved on ice and Atg7 (0.5 mM), Atg3 (5 mM), liposomes (25% PE, 25% PI, 45% PC and 5% PS) and lipidation buffer were added. In order to generate a control sample, pE3 was handled identical to the dephosphorylation reaction, however, no protein phosphatases were added at this stage. Samples were moved on ice and Atg7, Atg3, liposomes and lipidation buffer were added (as specified above). A mix of lambda phosphatase, PP2A Rts1 and protein phosphatase inhibitors was added to the sample matching the protein phosphatase and protein phosphatase inhibitor concentration of the desphosphorylated E3 sample. Atg8 lipidation reactions containing either phosphorylated or dephosphorylated E3 were started by adding wildtype Atg8 (10 mM). Atg8 lipidation was monitored as a function of time and reactions were stopped by addition of 6X Urea sample buffer. Samples were analyzed by 15% Urea-SDS-PAGE.
Atg8 charging assays Atg8$Atg7 and Atg8$Atg3 thioester formation was monitored by mixing Atg7 (0.5 mM) and Atg3 (5 mM) with TMR labeled Atg8 M1C/DR117 (20 mM) in the presence of either 50 nM wild-type Atg1 or catalytically inactive Atg1 D211A . Atg3/Atg7 and TMR-Atg8 M1C/DR117 were separately incubated with either wild-type Atg1 or Atg1 D211A and the charging reactions were started by addition of TMR-Atg8 M1C/DR117 to the Atg3/Atg7 mixture. Phosphorylation and charging reactions were incubated at 30 C in 20 mM HEPES-NaOH pH 7.4, 120 mM NaCl, 2 mM ATP, 10 mM MgCl 2 . Reactions were stopped by mixing 6 ul charging reaction with 8 ul 6X reducing agent free LDS sample buffer. Control samples were stopped by addition of 6X LDS loading dye containing 100 mM DTT. Samples were incubated at 50 C for 10 minutes and analyzed by SDS-PAGE. The resultant gels were imaged on a Typhoon scanner and subsequently stained with InstantBlue (Expedeon).

Atg3 discharge assays
Atg8, Atg3 and Atg7 were individually incubated with either 50 nM wild-type Atg1 or catalytically inactive Atg1 D211A before the charging reactions containing 10 mM Atg8, 5 mM Atg3-StrepII2x and 2 mM Atg7 were prepared. Atg3 charging was carried out in 8 mM HEPES-NaOH pH 7.4, 50 mM NaCl, 4 mM MgCl 2 and 0.8 mM ATP. Samples were incubated for 30 minutes and EDTA was added to a final concentration of 50 mM. Reactions were started by addition of liposomes (1 mg/ml; containing 55% PE, 30% DOPC and 15% PI) and stopped by mixing 6 mL sample with 8 mL 6X reducing agent free LDS sample buffer. Control samples were taken at time point zero by adding 6X LDS loading dye containing 100 mM DTT. Samples were incubated at 50 C for 10 minutes and analyzed by SDS-PAGE and Coomassie staining.
Pho8D60 assays ''YPD'' samples were prepared by growing yeast cells in nutrient-rich YPD medium at 30 C to an OD 600 of $1.2. ''SD-N'' samples were prepared by switching yeast cells grown in YPD medium to starvation medium (SD-N). Cells were washed twice with SD-N medium before growing them for another 4 hours in SD-N medium (unless specified otherwise). Cells were harvested at 4000 rpm. Pellets were washed with ice-cold water, spun again and resuspended in resuspension solution (0.85% NaCl, 1 mM PMSF). Cells were processed as described previously (Klionsky, 2007) and alkaline phosphatase activity was measured using an end-point spectrophotometric assay monitoring hydrolysis of p-nitrophenolphosphate (pNPP) to p-nitrophenol (pNP). The average and standard deviations were calculated based on at least three biological replicates.

Pulldown assays
For Strep, Myc and FLAG pulldown experiments proteins were immobilized using either Strep-Tactin Superflow Plus (QIAGEN), Antic-Myc Agarose (Thermo Fisher) or Anti-FLAG M2 affinity resin (Sigma). Proteins were added and incubated with the resin for 15 minutes at room temperature. Beads were washed three times with 15-20 bed volumes each. Bound proteins were eluted using either 2.5 mM desthiobiotin (Sigma), 500 mg/ml Myc peptide (GenScript) or 100 mg/ml 3X FLAG peptide (Sigma) and analyzed by SDS-PAGE.

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Liposome pelleting assays Atg8 (10 mM) was lipidated as described above. Atg8 containing liposomes were pelleted and washed prior to sample addition. Proteins were incubated with Atg8 containing liposomes for 30 minutes. Liposomes were washed three times using wash buffer (20 mM HEPES-NaOH pH 7.4, 120 mM NaCl and 2 mM DTT). Liposomes were resuspended in Urea containing SDS sample buffer, and samples were analyzed by SDS-PAGE.

Fluorescence microscopy
For fluorescence microscopy experiments yeast cells were either exponentially grown in YPD medium or starved for 4 hours in nitrogen starvation medium (SD-N). Cells grown in YPD were pelleted and resuspended in synthetic complete (SC) medium prior to imaging. Images were acquired in a temperature-controlled environment (30 C) on an inverted wide-field Nikon Eclipse Ti microscope. Images were taken with a 100x oil objective. Atg proteins were imaged in the GFP channel and z stacks were recorded. Images were prepared using Fiji.
For co-localization experiments yeast strains yAS_610 and yASC_842 were transformed with the pRS425-tagBFP-Ape1 plasmid (Sch€ utter et al., 2020;kindly provided by Martin Gr€ af), and grown to early log-phase in synthetic defined medium lacking leucine (SD-LEU) at 30 C. Cells were pelleted and resuspended in SD-LEU medium containing 250 mM CuSO 4 and grown for another 3 hours. Cells were pelleted and washed 3 times in starvation medium (SD-N) and grown for 2 h 30 min in SD-N before imaging. The image datasets were acquired using a wide-field Nikon Ti2 inverted microscope, equipped with a 100X/1.45 NA lens and a Prime 95B sCMOS camera (Teledyne Photometrics), controlled through Micro-Manager 2.0 software (Edelstein et al., 2014). Z stacks of the BFP, neonGreen, and transmitted light channels were acquired using selective band-pass filters, over a range of 10 mm, every 0.25 mm. Deconvolution of the fluorescence channels was obtained processing the datasets with scikit-image 0.18.1 python package (van der Walt et al., 2014), adopting the Richardson-Lucy algorithm, with 15 iterations.

Fluorescence recovery after photobleaching (FRAP) experiments
Atg11 deleted yeast expressing Atg13-neonGreen and either wild-type Atg1 or catalytically inactive Atg1 D211A (yAS_621 and yAS_554 respectively) were grown in YPD medium and switched to nitrogen starvation medium (SD-N). Cells were grown in SD-N medium for 3 hours before they were imaged using a Leica TSC SP8 confocal microscope. GFP was excited with an argon laser at 488 nm, and emission was recorded between 498 nm-758 nm. Cells were imaged with a 63 3 /1.40 oil objective and images were acquired every 10 s for one minute after photobleaching. Images for each time point were bleach corrected and the fluorescence intensity of the bleached area was compared to the initial intensity after background subtraction.

Mass spectrometry
After in vitro phosphorylating Atg proteins (2 mM) with Atg1 (50 nM), samples were TCA precipitated and the resultant pellets washed twice with ice-cold acetone and resuspended in ABC urea buffer (50 mM NH 4 HCO 3 , 8 M Urea). TCEP was added to a final concentration of 5 mM. After 30 minutes at room temperature, iodoacetamide was added to a final concentration of 10 mM and reactions were incubated for another 30 minutes in the dark. Samples were diluted to a final urea concentration of 6 M using ABC buffer (50 mM NH 4 HCO 3 )), before incubating for 4-5 hours at 37 C with Lysyl-endopeptidase (LysC; Wako) (1:100). Reactions were diluted to 2 M Urea and incubated with trypsin overnight shaking in the dark. Formic acid (FA) was added to a final concentration of 1% and samples were loaded on a pre-equilibrated C18 Sep-PAK column (Waters). Columns were washed with 2% acetonitrile (ACN) containing 0.1% FA and samples were eluted with 50% ACN containing 0.1% FA. Samples were dried in a SpeedVac, resuspended in 5% ACN containing 0.1% FA, sonicated and analyzed by mass spectrometry. The same procedure was followed for endogenous Atg3 and Atg12 containing samples with the only differences that samples were not TCA precipitated and only trypsin was used for protein digestion. C18 cleaned samples were phospho-enriched using the High-Select Fe-NTA Phosphopeptide Enrichment Kit (Thermo Fisher Scientific) following the manufacturer's instructions.
Mass spectrometry data were acquired in data dependent acquisition mode using a 5600 TripleTOF (Sciex) or LTQ-Orbitrap XL (Thermo Fisher Scientific) instrument for in vitro kinase reactions, or a TimsTOF Pro for endogenously purified Atg3 samples. For Tri-pleTOF acquisition peptides were separated using an Eksigent NanoLC Ultra nanoLC system using a 60 min gradient from 2%-35% (buffer A 0.1% (v/v) formic acid, 2% (v/v) acetonitrile, buffer B 0.1% (v/v) formic acid, 90% (v/v) acetonitrile) after direct injection onto a 20-cm PicoFrit emitter (New Objective) packed to 20 cm with Magic C18 AQ 3-mm 200-Å stationary phase. MS1 spectra were collected for 250 ms with nominal resolving power of 30,000. The 20 most intense precursors with charge state 2-5 were selected for fragmentation, and MS2 spectra were collected in the range 50-2,000 m/z for 100 ms with nominal resolving power of 15,000 (high sensitivity mode); precursor ions were excluded from reselection for 15 s. For LTQ-Orbitrap XL acquisition peptide separation was carried out by reversed phase on a Proxeon EASY-nLC II liquid chromatography system (Thermo Fisher Scientific). The reverse phase column (75 mm x 10 cm) was packed with Magic C18 AQ 3-mm 200-Å stationary phase. A linear gradient from 5% to 35% acetonitrile in 0.1% formic acid was run for 60 min at a flow rate of 300 nL/min. Data acquisition was set to obtain one high resolution MS scan in the Orbitrap (60,000 at 400 m/z) followed by six collision induced fragmentation (CID) MS/MS fragment ion spectra in the linear trap quadrupole (LTQ). Orbitrap charge state screening was enabled and ions with unassigned or single charge states were rejected. The dynamic exclusion window was set to 15 s and limited to 300 entries. The minimal precursor ion current to trigger CID and MS/MS scan was set to 150. The ion accumulation time was set to 500 ms (MS) and 250 ms (MS/MS) using a target setting ll OPEN ACCESS Article e9 Molecular Cell 81, 5066-5081.e1-e10, December 16, 2021 of 10 6 (MS) and 10 4 (MS/MS) ions. For TimsTOF Pro data acquisition peptide separation was carried on a nanoElute liquid chromatography system (Bruker) using a reversed phase nanoElute TEN (Bruker) 75mm x 10 cm column packed with 1.9 mm C18 ReproSil AQ beads maintained at 50 C using a linear gradient from 3% to 28% acetonitrile in 0.1% formic acid at 400 nl/min over 60 minutes. Data was acquired in ddaPASEF mode (Meier et al., 2018) using the vendor provided standard 1.1 s cycle time acquisition method (10 PASEF ramps per cycle, collision energy ramp 20-59 V).
MS/MS spectra were searched in MaxQuant v1.6.12.0 (Cox and Mann, 2008) against a yeast protein sequence database retrieved from UniProt (September 2020 -strain ATCC 204508 / S288c -taxon identifier -559292 -containing 6164 sequences) appended with common contaminants, affinity tag sequences, and decoys generated by protein sequence reversal. Search settings were set to fully-tryptic cleavage with 2 missed cleavages allowed, variable modification of oxidation allowed on methionine, phosphorylation on serine/threonine/tyrosine, acetylation on protein N terminus and fixed modification of carbamidomethylation on cysteines. All other search parameters were set to default for the respective instruments. False discovery rate was controlled at 1% at peptide and protein levels. For quantification we extracted MS1 level extracted Ion chromatograms using Skyline v20.2.1.404. Mapping of peptide identifications and phosphosite localizations was performed by importing MaxQuant search results into Skyline.

QUANTIFICATION AND STATISTICAL ANALYSIS
As indicated in the figure legends data are represented as average ± standard deviation.        Supplementary Figure 4. Phosphorylation of the Atg13 central region triggers Atg1-Atg13 complex dissociation.
A) The Atg1 D211A -Atg13 and Atg1 D211A -Atg13 S/T>A complex were incubated with the Atg17-Atg29-Atg31 subcomplex and with or without wild type Atg1 in the presence of ATP/Mg 2+ (I). Complexes were immobilized using StrepTactin Sepharose and StrepII 2x -Atg1 D211A and co-purifying proteins were eluted (E) and subsequently incubated with λ-PP (E+λ) prior to analysis by SDS-PAGE and Coomassie staining. B) The Atg13 S/T>A mutant does not dissociate from Atg1 D211A upon Atg1 mediated phosphorylation. The Atg1 D211A -Atg13 and Atg1 D211A -Atg13 S/T>A complex were incubated with or without substoichiometric amounts of wild type Atg1 in the presence of ATP/Mg 2+ (I). Complexes were immobilized using StrepTactin Sepharose and StrepII 2x -Atg1 D211A and co-purifying proteins were eluted (E) and subsequently incubated with λ-PP (E+λ) prior to analysis by SDS-PAGE and Coomassie staining. C) In vitro kinase assays studying phosphorylation of Atg3 and Atg12-Atg5-Atg16 (5 μM) by Atg1, Atg1-Atg13 or Atg1-Atg13 S/T>A (50 nM). Substrate phosphorylation was studied in a time dependent manner and individual time points were analysed by SDS-PAGE and autoradiography. Relative quantifications of substrate phosphorylation are depicted below with the average and standard deviation plotted for each time point. D) Fluorescence imaging of nitrogen starved S. cerevisiae cells expressing neonGreentagged wild type Atg13 or the Atg13 S/T>A mutant in the presence of either wild type Atg1 or catalytically inactive Atg1 (Atg1 D211A ). E) Fluorescence imaging of exponentially growing (YPD) or nitrogen starved (SD-N) S. cerevisiae cells expressing either Atg13-neonGreen or neonGreen-Atg17. Atg13-neonGreen was imaged in either wild type or atg11Δ, atg17Δ or atg11Δ atg17Δ deletion backgrounds and neonGreen-Atg17 was imaged in either wild type or atg11Δ, atg13Δ or atg11Δ atg13Δ deletion backgrounds. F) Fluorescence imaging of exponentially growing (YPD) or nitrogen starved (SD-N) S. cerevisiae cells expressing either neonGreen-Atg17 or Atg1-neonGreen in the presence of wild type Atg13 (WT) or the Atg13 S/T>A mutant. (D-F) Maximum intensity z-projections of deconvolved images are shown. Scale bar: 5 μm. G) Western blot quantification of the Cvt assay shown in Figure 3L (n=3).   Figure 5. Atg8 lipidation is inhibited by Atg1 dependent E3 phosphorylation. Figure 4 Supplementary Figure 5. Atg8 lipidation is inhibited by Atg1 dependent E3 phosphorylation. A) Atg7, Atg10, Atg12 and the Atg5-Atg16 complex were individually incubated with either wild type Atg1 or catalytically inactive Atg1 (Atg1 D211A ). Proteins were combined to initiate Atg12 conjugation to Atg5. Samples were taken at the indicated time points and analyzed by SDS-PAGE and Coomassie staining. B) SDS-PAGE comparison of Atg1 phosphorylated and non-phosphorylated Atg12-Atg5-Atg16 after anion exchange and size exclusion chromatography. Asterisk indicates co-purifying insect cell protein. C) Lipidation of Atg8 (Atg8 ΔR117 ) and Atg8 N (Atg8 S3A/T4A/S7A/ΔR117 ) are inhibited in the presence of Atg1 kinase activity. Full gel images of cropped images depicted in Figure 4B with neighbouring controls to confirm the inhibitory effect of Atg1 kinase activity on Atg8 and Atg8 N lipidation. D) Atg8 and Atg8 N were lipidated before incubating the reactions with increasing amounts of wild type Atg1 or catalytically inactive Atg1 D211A . The electrophoretic mobility shift of Atg8 and Atg8 N was monitored by Urea-SDS-PAGE and Sypro Ruby staining. E) ) In vitro lipidation kinetics of wild type Atg8 and Atg8 N . Samples were taken at the indicated time points and analyzed by Urea-SDS-PAGE and Sypro Ruby staining. The average Atg8-PE formation and standard deviation is plotted (n=3). F) The Pho8Δ60 assay was used to quantify bulk autophagy in atg8Δ strains or in cells expressing wild type Atg8 or the Nterminal alanine mutant Atg8 N . Cells were either exponentially grown in YPD medium or starved for 4 hours in nitrogen starvation medium (SD-N). Pho8Δ60 activity was measured in three independent experiments as described in 'Materials and Methods' and plotted as relative Pho8Δ60 activity with standard deviation. G) Schematic overview of the set up used to monitor Atg8 lipidation in the presence of either Atg1 phosphorylated or dephosphorylated E3 (pE3 and E3 respectively). Firstly, the E3 (Atg12-Atg5-Atg16-FLAG) was phosphorylated by Atg1 and subjected to a FLAG affinity purification and anion exchange chromatography step in order to minimize the Atg1 levels in the resultant pE3 preparation. To ensure that differences in Atg8 lipidation are not due to residual Atg1 kinase activity in the pE3 preparation the pE3 was dephosphorylated using lambda protein phosphatase (λ-PP) and recombinant PP2A Rts1 to act as the non-phosphorylated E3 control. The results and quantifications of the lipidation reactions are shown in Figure 4C. Liposomes used contained 25% PE, 45% PC, 5% PS and 25% PI. H) Wild type Atg12 and the phosphomimicking mutant, Atg12 S>D_ALL , were phosphorylated by Atg1 in a time dependent manner. Phosphorylation was analyzed by SDS-PAGE and auto-radiography. I) Atg8 lipidation was monitored in atg11Δ cells expressing either wild type Atg12 or the Atg12 mutants Atg12 S>D_ALL , Atg12 S>D_Ubl or Atg12 S>A_ALL . Cells deleted for ATG12 (atg12Δ) were used as a negative control. Cells were nitrogen starved for 18 hours and Atg8-PE formation was monitored by Western blotting using an anti-Atg8 antibody. J) Bulk autophagy was monitored in vivo using the Ape1 processing assay. Bulk autophagy dependent maturation of precursor Ape1 (prApe1) to its mature form (mApe1) was compared in atg11Δ cells expressing either wild type Atg12 or the phosphomimicking Atg12 S>D_N-term mutant. Cells were either exponentially grown in YPD medium or nitrogen starved for 5 or 18 hours. Ape1 processing was analyzed by Western blotting using an anti-Ape1 antibody. K) The cytoplasm-to-vacuole targeting (Cvt) pathway was monitored in S. cerevisiae cells expressing either wild type Atg12 or the phosphomimicking Atg12 S>D_N-term mutant. Cells were grown exponentially in nutrient-rich YPD medium and Ape1 processing was monitored by Western blotting. The conversion of precursor Ape1 (prApe1) to its mature form (mApe1) was quantified in the right panel (n=3). L) The Pho8Δ60 assay was used to quantify bulk autophagy in atg12Δ strains or in cells expressing wild type Atg12 or the phosphomimicking Atg12 S>D_N-term mutant. Cells were either exponentially grown in YPD medium or nitrogen starved for 2.5, 5, 7.5 or 18 hours. Alkaline phosphatase activity was measured (n=3) and plotted as relative Pho8Δ60 activity with standard deviation. M) E3 complex formation was compared between wild type Atg12 and the phosphomimicking Atg12 S>D_ALL mutant which has all detected Atg1 dependent in vitro phosphorylation sites mutated to aspartate. E3 complex formation was monitored in a time dependent manner after adding Atg12 or StrepII2x-FLAG-tagged Atg12 S>D_ALL (SF-Atg12) to Atg5-Atg16, Atg10, Atg7 and ATP/Mg 2+ containing reactions. Time points were analysed by SDS-PAGE and Coomassie staining.  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